Method

Defibrinated blood should be used, care being taken to ensure that sterility is maintained.

Incubate 1 ml or 2 ml volumes of blood in sterile 5 ml bottles. It is advisable to set up the samples in duplicate in case one has become infected, as indicated by gross lysis and change in colour.

After 24 h, if no infection is evident, pool the contents of the duplicate bottles after thoroughly mixing the sedimented red cells in the overlying serum and estimate the fragility as previously described.

Because the fragility may be markedly increased (Fig. 12.2), set up additional hypotonic solutions containing 7.0 g/l and 8.0 g/l NaCl. In addition, use a solution equivalent to 12.0 g/l NaCl because sometimes, as in HS, lysis may take place in 9.0 g/l NaCl. In this case, use the supernatant of the tube containing 12.0 g/l NaCl as the blank in the colorimetric estimation.

Figure 12.2 Osmotic fragility curves before and after incubating blood at 37°C for 24 h. Results are from patients suffering from the following: hereditary spherocytosis, pyruvate kinase deficiency and hereditary non-spherocytic haemolytic anaemia of undiagnosed type. The normal range is indicated by the unbroken lines.

The incubation fragility test is conveniently combined with the estimation of the amount of spontaneous autohaemolysis (p. 252).

Factors Affecting Osmotic Fragility Tests

In carrying out osmotic fragility tests by any method, three variables capable of markedly affecting the results must be controlled, quite apart from the accuracy with which the saline solutions have been made up. These are as follows:

A proportion of 1 volume of blood to 100 volumes of saline is chosen because the concentration of blood is so small that the effect of the plasma on the final tonicity of the suspension is negligible. When weak suspensions of blood in saline are used, it is necessary to control the pH of the hypotonic solutions and it is for this reason that phosphate buffer is added to the saline. Even so, small differences will be found between the fragility of venous blood and maximally aerated (i.e. oxygenated) blood. For the most accurate results, it is recommended that the blood should be mixed until bright red. Finally, it is ideal for tests to be carried out always at the same temperature, although for most purposes room temperature is sufficiently constant.

The extent of the effect of pH and temperature on osmotic fragility was well illustrated in the paper of Parpart et al.³ The effect of pH is more important: a shift of 0.1 of a pH unit is equivalent to altering the saline concentration by 0.1 g/l, the fragility of the red cells being increased by a decrease in pH. An increase in temperature decreases the fragility, an increase of 5°C being equivalent to an increase in saline concentration of about 0.1 g/l.

Lysis is virtually complete at the end of 30 min at 20°C and the hypotonic solutions may be centrifuged at the end of this time.

Further details of the factors that affect and control haemolysis of red cells in hypotonic solutions were given by Murphy.⁴

Recording the Results of Osmotic Fragility Tests

In the past, osmotic fragility most often has been expressed in terms of the highest concentration of saline at which lysis is just detectable (initial lysis or minimum resistance) and the highest concentration of saline in which lysis appears to be complete (complete lysis or maximum resistance). It is, however, useful also to record the concentration of saline causing 50% lysis (i.e. the median corpuscular fragility, MCF) and to inspect the entire fragility curve (Fig. 12.1). The findings in health are summarized in Table 12.1.

Table 12.1 Osmotic fragility in health (at 20°C and pH 7.4)

MCF, median corpuscular fragility.

Interpretation of Results

The osmotic fragility of freshly taken red cells reflects their ability to take up a certain amount of water before lysing. This is determined by their volume-to-surface area ratio. The ability of the normal red cell to withstand hypotonicity results from its biconcave shape, which allows the cell to increase its volume by about 70% before the surface membrane is stretched; once this limit is reached lysis occurs.⁶ Spherocytes have an increased volume-to-surface area ratio; their ability to take in water before stretching the surface membrane is thus more limited than normal and they are therefore particularly susceptible to osmotic lysis. The increase in osmotic fragility is a property of the spheroidal shape of the cell and is independent of the cause of the spherocytosis. Characteristically, osmotic fragility curves from patients with HS who have not been splenectomized show a ‘tail’ of very fragile cells (Fig. 12.3). When plotted on probability paper, the graph indicates two populations of cells: the very fragile and the normal or slightly fragile. After splenectomy the red cells are more homogeneous, the osmotic fragility curve indicating a more continuous spectrum of cells, from fragile to normal.

Figure 12.3 Osmotic fragility curves of three patients suffering from hereditary spherocytosis belonging to the same family (brother, sister and uncle [Le Ay]). The area between the unbroken lines represents the normal range.

Decreased osmotic fragility indicates the presence of unusually flattened red cells (leptocytes) in which the volume-to-surface area ratio is decreased. Such a change occurs in iron deficiency anaemia and thalassaemia in which the red cells with a low mean cell haemoglobin (MCH) and mean cell volume (MCV) are unusually resistant to osmotic lysis (Fig. 12.1). A simple one-tube osmotic fragility is a useful screening test for β thalassaemia and some haemoglobinopathies in countries with a high incidence of these abnormalities (p. 612). Reticulocytes and red cells from patients who have been splenectomized also tend to have a greater amount of membrane compared with normal cells and are osmotically resistant. In liver disease, target cells may be produced by passive accumulation of lipid and these cells, too, are resistant to osmotic lysis.⁷

The osmotic fragility of red cells after incubation for 24 h at 37°C is also a reflection of their volume-to-surface area ratio, but the factors that alter this ratio are more complicated than in fresh red cells. The increased osmotic fragility of normal red cells, which occurs after incubation (Fig. 12.2), is mainly caused by swelling of the cells associated with an accumulation of sodium that exceeds loss of potassium. Such cation exchange is determined by the membrane properties of the red cell, which control the passive flux of ions, and the metabolic competence of the cell, which determines the active pumping of cations against concentration gradients. During incubation for 24 h, the metabolism of the red cell becomes stressed and the pumping mechanisms tend to fail, one factor being a relative lack of glucose in the medium.

The osmotic fragility of red cells that have an abnormal membrane, such as those of HS and hereditary elliptocytosis (HE), increases abnormally after incubation (Fig. 12.2). Similar results occur in hereditary stomatocytosis.⁸ The results with red cells with a glycolytic deficiency, such as those of pyruvate kinase (PK) deficiency, are variable. In severe deficiencies, osmotic fragility may increase substantially (Fig. 12.2), but, in other cases, the fragility may decrease owing to a greater loss of potassium than gain of sodium. In thalassaemia major and minor, osmotic fragility is frequently markedly reduced after incubation, again owing to a marked loss of potassium.⁹ A similar, although usually less marked, change is seen in iron deficiency anaemia.

To summarize, measurement of red cell osmotic fragility provides a useful indication as to whether a patient’s red cells are normal because an abnormal result invariably indicates abnormality. The reverse is, however, not true (i.e. a result that is within the normal range does not mean that the red cells are normal). The findings in some important haemolytic anaemias are summarized in Table 12.2.

Table 12.2 Osmotic fragility in haemolytic anaemias: a summary

OF, osmotic fragility; HE, hereditary elliptocytosis; HS, hereditary spherocytosis; MCF, median corpuscular fragility; MCHC, mean cell haemoglobin concentration.

Principle

The osmotic fragility test lacks specificity and sensitivity and fails to detect atypical or mild HS. Moreover, it can be affected by factors unrelated to red cell cytoskeletal defects; for example, positive results may be obtained for red cells from patients who are pregnant or who have immune or other haemolytic anaemias or renal failure. The flow cytometric (dye-binding) test of King and colleagues¹⁰ measures the fluorescent intensity of intact red cells labelled with eosin-5-maleimide (EMA), which reacts covalently with Lys-430 on the first extracellular loop of Band 3 protein. The N-terminal cytoplasmic domain of Band 3 interacts with ankyrin and protein 4.2, which in turn crosslink with the spectrin-based cytoskeleton and stabilizes the membrane lipid bilayer.¹¹ Deficiency or abnormality of Band 3 may result in decreased fluorescence. This is seen in HS red cells but has also been observed in cases of South-east Asian ovalocytosis, congenital dyserythropoietic anaemia Type II and the stomatocytic variant, cryohydrocytosis. Blood samples in ethylenediaminetetra-acetic acid (EDTA) may be analysed for up to 48 h after collection provided they have been stored in the refrigerator.

Reagents

Method

Thaw a tube of stock EMA solution in the dark at room temperature and dilute with an equal volume of PBS to obtain a working solution of 0.5 mg/ml. Mix 5 μl of washed packed red cells with 25 μl of EMA working solution in a plastic microfuge tube. Set up control tubes from blood of normal individuals and perform all tests in duplicate. Leave a rack with the tubes in a cupboard in the dark at room temperature for 30 min. Mix and return to the cupboard for a further 30 min.

Then, spin the tubes in a bench-top microfuge for 5–10 s and remove the supernatant dye carefully with a fine-tip pipette.

Wash the labelled red cells three times with 500 μl PBS containing 0.5% BSA. The third wash should be colourless. If it is still pink, suggesting that traces of dye particle remain in the tube, discard the sample and repeat the cell labelling procedure.

Resuspend the packed red cells in 500 μl of the PBS–BSA wash solution. Transfer 100 μl of the cell suspension into a plastic flow cytometer tube and add 1.4 ml of wash solution. Keep the cell suspensions in the dark by wrapping in aluminium foil until use. Set the analyser thresholds (gate) for red blood cells and count each sample for a minimum of 15 000 events. Select the FITC (fluorescein isothiocyanate) channel and record the mean fluorescence intensity (MFI). Compare the test with the mean value of several control samples.

Interpretation of Results

Results should be expressed as a ratio of mean value of the test to control sample. Each laboratory should set the reference range and cut-off values for its own instrument. In our laboratory, a ratio of 0.8 or more is regarded as normal.

Principle

Glycerol present in a hypotonic buffered saline solution slows the rate of entry of water molecules into the red cells so that the time taken for lysis may be more conveniently measured. Like the osmotic fragility test, differentiation can be made between spherocytes and normal red cells.

Reagents

Method

Add 20 μl of whole blood, anticoagulated with EDTA, to 5.0 ml of PBS, pH 6.85. Mix the suspension carefully.

Transfer 1.0 ml to a standard 4 ml cuvette of a spectrometer equipped with a linear-logarithmic recorder. Fix the wavelength at 625 nm and start the recorder. Add 2.0 ml of the glycerol reagent rapidly to the cuvette with a 2 ml syringe or automatic pipette and mix well.

The rate of haemolysis is measured by the rate of fall of turbidity of the reaction mixture. The results are expressed as the time required for the optical density to fall to half the initial value (AGLT₅₀). The test can also be carried out using a colorimeter and stopwatch.

Results

Normal blood takes more than 30 min (1800 s) to reach the AGLT₅₀. The time taken is similar for blood from normal adults, newborn infants and cord samples. In patients with HS, the range of the AGLT₅₀ is 25–150 s. A short AGLT₅₀ may also be found in chronic renal failure, chronic leukaemias and autoimmune haemolytic anaemia; it also may be found in some pregnant women.¹⁴

Significance of the AGLT

The same principles apply as with the osmotic fragility test. Cells with a high volume-to-surface area ratio resist swelling for a shorter time than normal cells. This applies to all spherocytes, whether the spherocytosis is caused by HS or other mechanisms. The test is particularly useful in screening family members of patients with HS where morphological changes are too small to indicate clearly whether the disorder is present.

Principle

Whereas osmotic fragility may be abnormal in any condition where spherocytes occur, it has been suggested that cryohaemolysis is specific for HS.¹⁵ This appears to result from the fact that the latter is dependent on factors that are related to molecular defects of the red cell membrane rather than to changes in the surface area-to-volume ratio. The test can be carried out on EDTA blood up to 1 day old.

Reagent

Method¹⁵

Interpretation

Streichman et al.¹⁵ report the range of cryohaemolysis in normal subjects to be 3–15%, whereas in hereditary spherocytosis there is >20% lysis. However, it is recommended that individual laboratories establish their own reference values for the method. We have found that most normal samples give <3% lysis. Increased lysis is not exclusive to hereditary spherocytosis and may be observed in hereditary stomatocytosis.

Principle

Aliquots of blood are incubated both with and without sterile glucose solution at 37°C for 48 h. After this period, the amount of spontaneous haemolysis is measured colorimetrically.

Method

It is essential to use aseptic techniques in setting up the autohaemolysis test to maintain sterility throughout the incubation period.

Defibrinating Blood

Defibrinate blood, as described on p. 5.

Use sterile defibrinated blood and deliver four 1 ml or 2 ml samples into sterile 5 ml capped bottles. Retain a portion of the original sample; separate and store this as the preincubation serum.

Add to two of the bottles 50 or 100 μl of sterile 100 g/l glucose solution, so as to provide a concentration of glucose in the blood of at least 30 mmol/l. Make sure that the caps of the bottles are tightly closed and place the series of bottles in the incubator at 37°C. A sample from a known normal individual should be run in parallel as a control.

After 24 h, thoroughly mix the content by gentle swirling. After incubating for 48 h, inspect the samples for signs of infection, thoroughly mix again, then from each bottle remove a sample for the estimation of the packed cell volume (PCV) (by the microhaematocrit method) and haemoglobin (Hb) concentration and centrifuge the remainder to obtain the supernatant serum.

Estimate the spontaneous lysis by means of a colorimeter or a spectrometer at 540 nm.

As a rule, it is convenient to make a 1 in 10 dilution of the incubated serum in cyanide-ferricyanide (Drabkin’s) solution (p. 25), unless there is marked haemolysis, when a 1 in 25 or 1 in 50 dilution is more suitable. A corresponding dilution of the preincubation serum is used as a blank and a 1 in 100 or 1 in 200 dilution of the whole blood in Drabkin’s solution indicates the total amount of Hb present and serves as a standard.

Calculate the percentage lysis, allowing for the change in PCV resulting from the incubation as follows:

where R₀ = reading of diluted whole blood; R_t = reading of diluted serum at 48 h; B = reading of blank; PCV_t = packed cell volume at time t; D₀ = dilution of whole blood (e.g. 1 in 200 = 0.005); and D_t = dilution of serum (e.g. 1 in 10 = 0.1).

The reading at time t is multiplied by (1−PCV_t) so as to give the concentration that would be found if the liberated Hb was dissolved in whole blood (i.e. in both plasma and red cell compartments), not in the plasma compartment alone.

Normal Range of Autohaemolysis

Lysis at 48 h: without added glucose, 0.2–2.0%; with added glucose, 0–0.9%.

The results obtained are sensitive to slight differences in technique and each laboratory should use a carefully standardized procedure and establish its own normal range. If the amount of liberated Hb is small, it is more accurate (although more time consuming) to measure lysis by a chemical method rather than by a direct spectrometric method (p. 231). It can also be measured directly by a simple and rapid procedure with a HemoCue Plasma/Low Hb system.¹⁷

Significance of Increased Autohaemolysis

Little or no lysis takes place when normal blood is incubated for 24 h under sterile conditions and the amount present after 48 h is small.¹⁶ If glucose is added so that it is present throughout the incubation, the development of lysis is markedly slowed. The amount of autohaemolysis that occurs after 48 h with and without glucose is determined by the properties of the membrane and the metabolic competence of the red cell. In membrane disorders such as HS, the rate of glucose consumption is increased to compensate for an increased cation leak through the membrane.⁸ During the 48-h incubation, glucose is therefore used up relatively rapidly so that energy production fails more quickly than normal unless glucose is added. This is one factor that contributes to the increased rate of autohaemolysis in HS. Usually, but not always, the addition of glucose to the blood decreases the rate of autohaemolysis in HS. This was referred to as Type 1 autohaemolysis.¹⁶ When the utilization of glucose via the glycolytic pathway is impaired, as in PK deficiency, the rate of autohaemolysis at 48 h is usually increased but glucose fails to correct or may even aggravate lysis (Type 2 autohaemolysis).⁸ Although a similar result may be seen in severe HS (Type B), in the absence of spherocytosis failure of glucose to diminish autohaemolysis is a strong indication of a glycolytic block. Blood from patients with G6PD deficiency or other disorders of the pentose phosphate pathway may undergo a slight increase in autohaemolysis (without additional glucose), which is corrected by the addition of glucose. Commonly, the result is normal, but examination of the incubated blood may show an increase in methaemoglobin (Hi) (discussed later). Not all glycolytic enzyme deficiencies give a Type 2 reaction so that a Type 1 result does not exclude the possibility of such a defect.

In the acquired haemolytic anaemias, the results of the autohaemolysis test are variable and generally not very helpful in diagnosis. In the autoimmune haemolytic anaemias, lysis may be increased in the absence of additional glucose but the effect of added glucose is unpredictable. In paroxysmal nocturnal haemoglobinuria (PNH), the autohaemolysis of aerated defibrinated blood is usually normal.

Autohaemolysis may be increased in haemolytic anaemias caused by oxidant drugs or when there are defects in the reducing power of the red cell. Heinz bodies, Hi or both will be detectable at the end of incubation. Normally, red cells produce <4% Hi after 48 h incubation and Heinz bodies are not seen. Red cells containing an unstable Hb also contain Heinz bodies at the end of the incubation period and increased amounts of Hi.

The nucleosides adenosine, guanosine and inosine, like glucose, diminish the rate of autohaemolysis when added to blood. Remarkably, adenosine triphosphate (ATP) strikingly retards haemolysis in PK deficiency, although glucose itself is ineffective.¹⁸ ATP does not pass the red cell membrane.

The autohaemolysis test lacks specificity. This has drawn much criticism on the test, including the suggestion that it has no place in the screening of blood for inherited defects.¹⁹ The best way to detect metabolic defects in red cells is undoubtedly to measure glucose consumption, lactate production and the contribution to metabolism of the pentose phosphate pathway. These measurements are, unfortunately, difficult and are likely to be undertaken only by specialized laboratories. The autohaemolysis test does provide some information about the metabolic competence of the red cells and helps to distinguish membrane defects from enzyme defects.

In summary, we feel that the autohaemolysis test is still useful in the investigation of patients who have or who may have chronic haemolytic anaemia for the following reasons:

Thus, in our experience, a combination of red cell morphology with the results of the autohaemolysis tests makes it possible to differentiate membrane abnormalities from enzyme deficiencies in the vast majority of cases.

Principle

NADPH, generated by G6PD present in a lysate of blood cells, fluoresces under long-wave ultraviolet (UV) light. In G6PD deficiency, there is an inability to produce sufficient NADPH; this results in a lack of fluorescence.

Reagents

Method

Thaw out sufficient tubes to set up test and controls. Allow reagents to reach room temperature before use.

Add 10 μl of whole blood (EDTA, heparin, ACD [acid–citrate–dextrose] or CPD [citrate–phosphate–dextrose]) to 100 μl volumes of the reagent mixture and keep at room temperature (15–25°C).

Place 10 μl of the reaction mixture on a Whatman No. 1 filter paper at the beginning of the reaction and again after 5–10 min. A shorter interval may be appropriate at a high ambient temperature (c 25–30°C). Allow to air dry thoroughly before examining the spots under UV light. Record whether fluorescence is present (+) or absent (−). Always set up samples of normal blood and known G6PD-deficiency blood in parallel.

If the samples are to be collected away from the laboratory or where delay is envisaged (e.g. during population screening) place about 10 μl of blood on Whatman No. 1 filter paper and allow it to dry. Cut out the disc of dried blood in the laboratory and add it to the reaction mixture. A sample of normal blood should always be tested as a positive fluorescence (i.e. normal) control.

The test can be carried out on blood stored in ACD (provided it is sterile) for up to 21 days at 4°C and for about 5 days at room temperature.

Interpretation

Fluorescence is produced by NADPH formed from NADP⁺ in the presence of G6PD. Some of the NADPH produced is oxidized by GSSG, but this reaction, catalysed by glutathione reductase, is normally slower than the rate of NADPH production. Red cells with <20% of normal G6PD activity do not cause detectable fluorescence.

Like all screening tests, this method is useful when large numbers of samples are to be tested, but the result must be interpreted with caution in an individual patient. The main causes of erroneous interpretations are as follows:

Although it is possible to correct for either or both of these contingencies, if in doubt, it is best to proceed directly to a quantitative enzyme assay (discussed later).

The test is meant to give only a + or − (normal or deficient) result by comparison with the controls and it does not make sense to grade by eye the intensity of fluorescence. If a control G6PD-deficient sample is not available, the appearance of the ‘zero time’ spot can be used for reference. The threshold for a ‘deficient’ result can be worked out by making dilutions of a normal blood sample in saline and is best set by regarding as deficient the fluorescence obtained when G6PD activity is 20% of normal or less (corresponding to a 1 in 5 dilution of normal blood). This means that very mildly deficient variants, and a substantial proportion of heterozygotes (see below), will be missed. However, clinically important haemolysis is unlikely to occur in subjects who have more than 20% G6PD activity and therefore this seems an appropriate (although arbitrary) threshold for a diagnostic laboratory. Because the test depends on visual inspection, it is best to select the time of incubation in relation to ambient temperature in preliminary trials. NADPH production is a cumulative process. Therefore, given enough time, a G6PD-deficient sample will fluoresce. The time allowed for the reaction should be one at which the contrast in fluorescence between a G6PD-normal and a G6PD-deficient sample is maximal.

Principle

Sodium nitrite converts Hb to Hi. When no methylene blue is added, methaemoglobin persists, but incubation of the samples with methylene blue allows stimulation of the pentose phosphate pathway in subjects with normal G6PD levels. The Hi is reduced during the incubation period. In G6PD-deficient subjects, the block in the pentose phosphate pathway prevents this reduction.²⁵

Reagents

Method

Use anticoagulated blood (EDTA or ACD) and test the samples preferably within 1 h of collection if left on the bench or within 6 h if kept at 4°C. Blood in ACD can be stored for up to 1 week but will be unsatisfactory if there is any haemolysis. With blood from patients who are severely anaemic, adjust the PCV to 0.40 ± 0.05.

Add 2 ml of blood to the tube containing 0.2 ml of the combined reagent either freshly prepared or dried. Close the tube with a stopper and gently mix the contents by inverting it 15 times.

Prepare control tubes by adding 2 ml of blood to a similar tube without reagents (normal reference tube) and to a tube containing 0.1 ml of sodium nitrite–dextrose mixture without methylene blue (‘deficient’ reference tube).

Incubate the samples at 37°C for 90 min. If the blood has been heparinized, incubation should be continued for 3 h.

After the incubation, pipette 0.1 ml volumes from the test sample, the normal reference tube and the deficient reference tube into 10 ml of water in separate, clear glass test tubes of identical diameter. Mix the contents gently. Compare the colours in the different tubes (see below).

Interpretation

Normal blood yields a colour similar to that in the normal reference tube (i.e. a clear red). Blood from deficient subjects gives a brown colour similar to that in the deficient reference tube. Heterozygotes give intermediate reactions.

Although this method takes longer than the fluorescent test, its advantages include the fact that it is extremely inexpensive and that the only equipment required is a waterbath. In addition, the test can be complemented by cytochemical analysis that lends itself to detecting G6PD deficiency in patients with reticulocytosis and in heterozygotes.

Test Kits

Several commercial kits are available for detection of G6PD deficiency. A fluorescent spot test (Trinity Biotech 203-A) and a test based on reduction of the dye dichloroindophenol to a colourless state in the presence of phenazine methosulphate (Trinity Biotech 400) are available commercially.*

The Quantase kit^† is a photometric method for use on whole blood or dried blood spots; NADPH produced by oxidation of G6P to 6PG is measured by an increase in absorption at 340 nm.

Each test or batch of tests should include a normal and a G6PD-deficient sample. Sheep blood is a useful source of naturally deficient blood. Where possible, participation in an external quality assessment (or proficiency testing) scheme is also recommended.

Reagents

Method

Venous blood collected into ACD should be used. The test should be carried out within 8 h of collection and the blood should be kept at 4°C until it is tested. Centrifuge the blood at 4°C for 20 min at 1200–1500 g.²⁸

Discard the supernatant and add 0.5 ml of the packed red cells to 9 ml of 9 g/l NaCl and 0.5 ml of sodium nitrite solution contained in a 15 ml glass centrifuge tube. Incubate at 37°C for 20 min. Centrifuge at 4°C for 15 min at approx. 500 g, then discard the supernatant fluid without disturbing the buffy coat and uppermost layer of red cells. Wash the cells three times in cold saline. After the last washing, remove the buffy coat, mix the packed cells well and transfer 50 μl to a glass tube containing 1 ml of the incubation medium. Incubate the suspension undisturbed at 37°C for 30 min. Then add 0.2 ml of MTT solution, shake gently and incubate at 37°C for 1 h. Resuspend the cells thoroughly. Place one drop adjacent to one drop of hypotonic saline on a glass slide, mix the drops thoroughly and cover with a coverglass.

Examine the red cells with an oil-immersion objective, noting the presence of formazan granules (Fig. 12.5).

Figure 12.5 Cytochemical demonstration of G6PD (glucose-6-phosphate dehydrogenase). Normal blood; positive reaction with formazan granules in the red cells.

Interpretation

When G6PD activity is normal, all the red cells are stained. In G6PD hemizygotes, the majority of the red cells are unstained. In heterozygotes, mosaicism is usually seen; usually 40–60% of the cells are unstained, but the proportion may be much less and in extreme cases only as few as 2–3% may be unstained.

Principle

The nucleotide pool of normal red cells consists largely (>96%) of purine (adenine and small amounts of guanine) derivatives. The levels of cytidine and uridine are normally extremely low. However, in P5N-deficient cells, more than 50% of this pool consists of pyrimidine nucleotides.

In acidic solutions, cytidine nucleotides have an absorbance maximum at approx. 280 nm, whereas adenine, guanine and uridine nucleotides absorb maximally at 260 nm. The ratio of absorbance at 260 nm to absorbance at 280 nm reflects the relative abundance of cytidine nucleotides; the absorbance ratio is lower when pyrimidine derivatives are increased.

Reagents

Method

For sample preparation, centrifuge blood freshly collected in EDTA at 1200 g for 5 min, remove the plasma and wash the cells three times with ice-cold 9 g/l NaCl solution. Add 1 ml of a 50% suspension of the washed red cells to 4 ml of ice-cold 4% perchloric acid (PCA) solution and then shake vigorously for 30 s. Transfer the clear supernatant obtained after centrifugation at 1200 g for 15 min to a small test tube. Prepare a sham extract by adding 1 ml of 9 g/l NaCl to 4 ml of 4% PCA solution.³⁵

Add 500 μl of water and 300 μl of 1 mol/l glycine buffer to each of two cuvettes. To correct for optical differences between the cuvettes, read the sample cuvette against the blank at 260 and at 280 nm, giving readings B²⁶⁰ and B²⁸⁰. Add 200 μl of the red cell extract to the sample cuvette and 200 μl of the sham extract to the blank cuvette. With the spectrometer zeroed at 260 nm on the blank cuvette, read the sample cuvette to obtain the value S²⁶⁰. Repeat the process at 280 nm to obtain the reading S²⁸⁰.

The A²⁶⁰/A²⁸⁰ absorbance ratio (R) is calculated by subtracting the cuvette blank readings (positive or negative) at 260 and 280 nm from the readings obtained on the red cell extract when blanked against the sham extract:

Interpretation

The A²⁶⁰/A²⁸⁰ absorbance ratio of freshly collected washed red cells has been reported to be 3.11 ± 0.41 (mean ± SD). Absorbance ratios of <2.29 imply that the concentration of cytidine nucleotide is increased and suggest a reduced level of P5N. Selective accumulation of pyrimidines owing to putative defect in CDP choline phosphotransferase has been reported in rare patients with a disorder that resembles P5N deficiency characterized by haemolytic anaemia and basophilic stippling.

Samples showing a significantly reduced absorbance ratio should have a specific assay for P5N carried out.³⁶ This is likely to require referral to a reference laboratory where a nucleotide profile may also be undertaken. Nucleotide extraction followed by radiolabelling and separation by high-performance liquid chromatography can be performed. The nucleotides have characteristic UV absorption spectra and retention times, which permit subsequent radiodetection and quantification.

Collection of Blood Samples

Blood samples may be anticoagulated with heparin (10 iu/ml blood), EDTA (1.5 mg/ml blood) or ACD (for formulae and volumes see below). In any of these anticoagulants, all normal enzymes are stable for 6 days (and most for 20 days) at 4°C and for 24 h at 25°C. However, enzyme variants in samples from patients may be less stable. Therefore, we recommend that ACD is used as anticoagulant and that the samples are tested promptly. Ideally, samples of blood should be transferred to central laboratories in tubes surrounded by wet ice at 4°C. Frozen samples are unsuitable because the cells are lysed by freezing. Further details of enzyme stability were given by Beutler.²¹ Approx. 1 ml of blood is required for each enzyme assay.

Preparation of Acid–Citrate–Dextrose (ACD) Solution – NIH-A

Sterilize the solution by autoclaving at 121°C for 15 min. Its pH is 5.4. For use, add 10 volumes of blood to 1.5 volumes of solution.

Separation of Red Cells from Blood Samples

Leucocytes and platelets generally have higher enzyme activities than red cells. Moreover, with many enzyme deficiencies, notably PK deficiency, the decrease in enzyme activity may be much less pronounced in leucocytes and platelets than in red cells or it even may be absent. It is therefore necessary to prepare red cells which are as free from contamination as possible. Various methods are suitable (see ICSH);³⁷ two are described in the following.

Washing the Red Cells

Centrifuge the anticoagulated blood at 1200–1500 g for 5 min and remove the plasma together with the buffy coat layer.

Resuspend the cells in 9 g/l NaCl (saline) and repeat the procedure three times. This will remove about 80–90% of the leucocytes.

This simple method is adequate in most instances when more complicated manoeuvres are impractical, but it has the disadvantage that some of the reticulocytes and young red cells are lost together with the buffy coat. In addition, the remaining leucocytes may still be sufficient to cause misleading results – for instance, in PK deficiency. Therefore, ideally the following method should be adopted.

Filtration through Microcrystalline Cellulose Mixtures

Pure red cell suspensions can be made from whole blood by filtering the blood through a mixed bed of microcrystalline cellulose (mean size 50 μm) and α-cellulose. Mix approx. 0.5 g of each type of cellulose with 20 ml of ice-cold saline; this gives sufficient slurry for 3–5 columns. The barrel of a 5 ml syringe is used as a column. The outlet of the syringe is blocked with absorbent cotton wool, equal in volume to the 1 ml mark on the barrel. Pour the well-shaken slurry into the column to give a bed volume of 1–2 ml after the saline has run through. Wash the bed with 5 ml of saline to remove any ‘fines.’ When the saline has run through, pipette 1–2 ml of whole blood onto the column, taking care not to disturb the bed. Collect the filtrate, and once the blood has completely run into the bed, wash the column through with 5–7 ml of saline. The column should be made freshly for each batch of enzyme assays and used promptly.

By this method, about 99% of the leucocytes and about 90% of the platelets are removed. About 97% of the red cells are recovered and reticulocytes are not removed selectively. The procedure should not alter the age or size of distribution of the recovered red cells compared to native blood. This should be checked with each new batch of cellulose by counting reticulocytes.

Wash the cells collected from the column twice in 10 volumes of ice-cold saline and finally resuspend them in the saline to give a 50% suspension.

Determine the Hb and/or red cell count in a sample of the suspension.

Preparation of Haemolysate

Mix 1 volume of the washed or filtered suspension with 9 volumes of lysing solution consisting of 2.7 mmol/l EDTA, pH 7.0, and 0.7 mmol/l 2-mercaptoethanol (100 mg of EDTA disodium salt and 5 μl of 2-mercaptoethanol in 100 ml of water); adjust the pH to 7.0 with HCl or NaOH.

Ensure complete lysis by freeze-thawing. Rapid freezing is achieved using a dry-ice acetone bath or methanol that has been cooled to −20°C. Thawing is achieved in a waterbath at 25°C or simply in water at room temperature. Usually the haemolysate is ready for use without further centrifugation, but a 1-min spin in a microfuge is recommended to remove any turbidity (this may be unsuitable for some red cell enzymes that are stroma-bound). Dilutions, when necessary, are carried out in the lysing solution. The haemolysate should be prepared freshly for each batch of enzyme assays. Most enzymes in haemolysates are stable for 8 h at 0°C, but it is best to carry out assays immediately. G6PD is one of the least stable enzymes in this haemolysate and its assay should be conducted within 1 or 2 h of the lysate being prepared. The storing of frozen cells or haemolysates is not recommended; it is preferable to store whole blood in ACD.

Control Samples

Control samples should always be assayed at the same time as the test samples even when a normal range for the various enzymes has been established.

Take the control samples of blood at the same time as the test samples and treat them in the same way. When receiving samples from outside sources, always ask for a normal ‘shipment control’ to be included.

Reaction Buffer

The ICSH recommendation is for a Tris-HCl/EDTA buffer that is appropriate for all the common enzyme assays. The buffer consists of 1 mol/l Tris-HCl and 5 mmol/l Na₂EDTA, the pH being adjusted to 8.0 with HCl.

Dissolve 12.11 g of Tris (hydroxymethyl) methylamine and 168 mg of Na₂EDTA in water; adjust the pH to 8.0 with 1 mol/l HCl and bring the volume to 100 ml at 25°C.

Only two assays will be described in detail – those for G6PD and PK. However, the principles of these assays apply to all other enzyme assays. The assays are carried out in a spectrometer at a wavelength of 340 nm unless otherwise indicated. A final reaction mixture of 1.0 ml (or 3.0 ml) is suitable, the quantities given in the text being for 1.0 ml reaction mixtures unless otherwise stated. All dilutions of auxiliary enzymes are made in the lysing solution and all working materials should be kept in an icebath until ready for use. The assays are carried out at a controlled temperature, 30°C being the most appropriate. Cuvettes loaded with the assay reagents should be preincubated at this temperature for 10 min before starting the reaction. In most cases, the reaction is started by the addition of substrate. Many spectrometers have a built-in or attached recorder, by which the absorbance changes can be conveniently measured. If no recorder is available, visual readings should be made every 60 s. In any case, the reaction should be followed for 5–10 min and it is essential to ensure that during this time the change in absorbance is linear with time.

Method

The assays are carried out at 30°C; the cuvettes containing the first four reagents and water are incubated for 10 min before starting the reaction by adding the substrate, as shown in Table 12.4. Commercial kits are also available.*

Table 12.4 Glucose-6-phosphate dehydrogenase assay

EDTA, ethylenediaminetetra-acetic acid; NADP, nicotinamide adenine dinucleotide phosphate; G6P, glucose-6-phosphate.

The change in absorbance following the addition of the substrate is measured over the first 5 min of the reaction. The value of the blank is subtracted from the test reaction, either automatically or by calculation.

Calculation of Enzyme Activity

The activities of the enzymes in the haemolysate are calculated from the initial rate of change of NADPH accumulation:

where 6.22 is the mmol extinction coefficient of NADPH at 340 nm and 10³ is the factor appropriate for the dilutions in the reaction mixture. Results are expressed per 10¹⁰ red cells, per ml red cells or per g Hb by reference to the respective values obtained with the washed red cell suspension. However, the ICSH recommendation is to express values per g Hb and it is ideal to determine the Hb concentration of the haemolysate directly. When doing this, use a haemolysate to Drabkin’s solution ratio of 1:25.

G6PD is very stable and, with most variants, venous blood may be stored in ACD for up to 3 weeks at 4°C without loss of activity.

Some enzyme-deficient variants lose activity more rapidly and this will cause deficiency to appear more severe than it is. Therefore, for diagnostic purposes, a delay in assaying well-conserved samples should not be a deterrent.

Normal Values

The normal range for G6PD activity should be determined in each laboratory. If the ICSH method is used, values should not differ widely from the given values. Results are expressed in enzyme units (eu), which are the μ moles (μmol) of substrate converted per min.

For adults, these values are 8.83 ± 1.59 eu/g Hb at 30°C. However, newborns and infants may have enzyme activity that deviates appreciably from the adult value.^37,38 In one study, the newborn mean activity was about 150% of the adult mean.³⁹

Interpretation of Results

In assessing the clinical relevance of a G6PD assay result, three important facts must be kept in mind:

In practice, the following notes may be useful:

Method

The preparation of haemolysate, buffer and lysing solution is exactly the same as for the G6PD assay. In the PK assay it is particularly important to remove as many contaminating leucocytes and platelets as possible because these cells may be unaffected by a deficiency affecting the red cells and may have high activities of PK. The principle of the assay is as follows:

The pyruvate so formed is reduced to lactate in a reaction catalysed by lactate dehydrogenase (LDH) with the conversion of NADH (reduced form of nicotinamide adenine dinucleotide) to NAD:

To ensure that this secondary reaction is not rate limiting, LDH is added in excess to the reaction mixture and the PK activity is measured by the rate of fall of absorbance at 340 nm.

The reaction conditions are established in a 1 ml cuvette at 30°C by adding all the reagents shown in Table 12.6, except the substrate PEP, to the cuvette and incubating them at 30°C for 10 min before starting the reaction by the addition of the PEP.

The amounts to be added for low-substrate conditions are also shown in Table 12.6.

The change in absorbance (A) is measured over the first 5 min and the activity of the enzyme in micromoles of NADH reduced/min/ml haemolysate is calculated as follows:

where 6.22 is the millimolar extinction coefficient of NADH at 340 nm.

Express results as for G6PD.

A blank assay should be carried out to be certain that the LDH is free of PK activity. Use the 2-mercapto-ethanol-EDTA stabilizing solution (p. 261) in place of haemolysate for both the blank and system mixtures. If no change in absorbance is observed, indicating that the LDH is free of contaminating PK, it is unnecessary to recheck on subsequent assays. Otherwise, the blank rate must be subtracted in computing the true enzyme activity each time.

Normal Values

As with all enzyme assays, a normal range should be determined for each laboratory. Values should however, not be widely different between laboratories if the ICSH methods are used. The normal range of PK activity at 30°C is 10.3 ± 2 eu/g Hb. At a low-substrate concentration, the normal activity is 15 ± 3% of that at the high-substrate concentration. Mean neonatal value is about 140% that of adults.³⁹

Interpretation of Results

PK, like G6PD, is an age-dependent red cell enzyme. But unlike G6PD deficiency, PK deficiency is usually associated with chronic haemolysis. Therefore patients in whom PK deficiency is suspected almost invariably have a reticulocytosis and if their PK level is below the normal range, they can be considered to be PK deficient. Thus, once the technique and normal values are well established in a laboratory, and provided controls are always included, the main problem is of underdiagnosis rather than of overdiagnosis of PK deficiency. One additional way to pick up abnormal variants has been included in the method recommended (i.e. the use of low-substrate concentrations). Even so, PK deficiency may be missed because marked reticulocytosis may increase PK activity significantly. This means that a PK activity in the normal range in the presence of a marked reticulocytosis is highly suspicious of inherited PK deficiency (because with reticulocytosis the activity ought to be higher than normal). In such cases, the importance of family studies cannot be overemphasized. Heterozygotes have about 50% of the normal PK activity, sometimes less, but they do not suffer from haemolysis. Therefore, the heterozygous parents of a patient may have a red cell PK activity lower than that of their homozygous PK-deficient offspring; this finding may clinch the diagnosis. In this context, assay of an alternative red cell age-dependent enzyme (e.g. G6PD or hexokinase) may be a useful aid to interpretation.

Principle

The method described is based on the development of a yellow colour when 5,5′-dithiobis (2-nitrobenzoic acid) (Ellman’s reagent, DTNB) is added to sulphydryl compounds. The colour that develops is fairly stable for about 10 min and the reaction is little affected by variation in temperature.

The reaction is read at 412 nm. GSH in red cells is relatively stable and venous blood samples anticoagulated with ACD maintain GSH levels for up to 3 weeks at 4°C. GSH is slowly oxidized in solution, so only fresh lysates should be used for the assay.

Reagents

Method

Add 0.2 ml of well-mixed, anticoagulated blood of which the PCV, red cell count and Hb have been determined, to 1.8 ml of lysing solution and allow to stand at room temperature for no more than 5 min for lysis to be completed, because glutathione oxidizes very rapidly when exposed to air.

Add 3 ml of precipitating solution, mix the solution well and allow to stand for a further 5 min.

After re-mixing, filter through a single-thickness Whatman No. 42 filter paper.

Add 1 ml of clear filtrate to 4 ml of freshly made Na₂HPO₄ solution. Record the absorbance at 412 nm (A₁). Then add 0.5 ml of the DTNB reagent and mix well by inversion.

The colour develops rapidly but begins to fade after about 10 min so the second reading should be within this period. Record the absorbance at 412 nm (A₂) and calculate the change in absorbance (ΔA⁴¹²).

A reagent blank is made using saline or plasma instead of whole blood.

If assays are carried out frequently, it is not necessary to construct standard curves for each batch. They are, however, essential initially to calibrate the apparatus used and should be done regularly to check the suitability of the reagents. Suitable dilutions of GSH are achieved by substituting 5, 10, 20 and 40 μl of the 1.62 mmol/l stock solution (make up to 0.2 ml with lysing solution) for the blood in the reaction.

Calculation

Calculation of GSH Concentration

The amount of GSH in the cuvette sample (GSH_c) is given by the following:

The concentration of GSH in the whole blood sample is as follows:

The unit is often expressed in terms of mg/dl of red cells. The molecular weight of GSH is 307. Thus, GSH in mg/dl packed red cells is given by the following:

Normal Range

The normal range may be expressed in a number of ways (e.g. 6.57 ± 1.04 µmol/g Hb or 223 ± 35 µmol (or 69 ± 11 mg)/dl packed red cells). Neonatal mean value is about 150% that of adults.³⁹

Significance

Glutathione replenishment in mature red cells is accomplished through the consecutive action of two enzymes: γ-glutamylcysteine synthetase and glutathione synthetase. Although very rare, hereditary deficiency of either enzyme virtually abolishes the synthesis of GSH. The deficient cells are very prone to oxidative destruction and are short lived, resulting in a non-spherocytic haemolytic anaemia.

Increases in GSH have been described in various conditions such as dyserythropoiesis, primary myelofibrosis, pyrimidine 5′-nucleotidase deficiency and other rare congenital haemolytic anaemias of unknown aetiology.

Principle

In normal subjects, incubation of red cells with the oxidizing drug acetylphenylhydrazine has little effect on the GSH content because its oxidation is reversed by glutathione reductase, which in turn relies on G6PD for a supply to NADPH. Therefore, in subjects who are G6PD deficient, the stability of GSH is significantly reduced.

Reagents

Method

Venous blood, anticoagulated with EDTA, heparin or ACD, may be used; it may be freshly collected or previously stored at 4°C for up to 1 week.

Add 1 ml to a tube containing acetylphenylhydrazine and place another 1 ml in a similar tube not containing the chemical. Invert the tubes several times and then incubate them at 37°C.

After 1 h, mix the contents of the tubes once more and incubate the tubes for a further 1 h. At the end of this time determine and compare the GSH concentration in the test sample and in the control sample.

Interpretation

In normal adult subjects, red cell GSH is lowered by not more than 20% by incubation with acetylphenylhydrazine. In subjects who are G6PD deficient, it is lowered by more than this: in heterozygotes (females), the fall may amount to about 50%, whereas in hemizygotes (males) the fall is often much greater and almost all may be lost.

The test is not specific for G6PD deficiency and other rare defects of the pentose phosphate pathway may give abnormal results.

Glutathione and Glutathione Stability in Infants

During the first few days after birth, the red cells have a normal or high content of GSH. On the addition of acetylphenylhydrazine, the GSH is unstable in both normal infants and infants who are G6PD deficient. In normal infants, however, the instability can be corrected by the addition of glucose and, by the time the normal infant is 3–4 days old, the cells behave like adult cells.^49,50

Principle

2,3-DPG is hydrolysed to 3PG by the phosphatase activity of PGM stimulated by glycolate-2-phosphate. This reaction is linked to the conversion of NADH to NAD by glyceraldehyde-3-phosphate dehydrogenase (Ga3PD) and phosphoglycerate kinase (PGK):

The fail in absorbance at 340 nm, as NADH is oxidized, is measured.

Reagents

Method

Freshly drawn blood in EDTA or heparin may be used. If there is an unavoidable delay in starting the assay, blood (4 volumes) should be added to CPD anticoagulant (1 volume) and stored at 4°C. A control blood sample should be taken at the same time.

2,3-DPG levels are stable for 48 h if the blood is stored in this way. The Hb, red cell count and PCV should be measured on part of the sample. It is not necessary to remove leucocytes or platelets.

Reaction

Deliver the reagents into a silica or high-quality glass cuvette, with a 1 cm light path. The quantities in Table 12.7 are for a 4 ml cuvette:

Table 12.7 Quantities for a 4 ml cuvette

Warm the mixtures at 30°C for 10 min and record the absorbance of both test and blank mixtures at 340 nm. Then start the reaction by the addition of 100 μl of glycolate-2-phosphate.

Remeasure the absorbance (at 35 min) of the test and blank mixtures on completion of the reaction.

Make further measurements after a further 5 min to make sure the reaction is complete.

Only one blank is required for each batch of test samples.

Calculation

2,3-DPG (µmol/ml blood)

where 3.10 = the volume of reaction mixture, 6.22 = mmolar extinction coefficient of NADH at 340 nm and 16 = dilution of original blood sample (1 ml in 3.0 ml of TCA, 0.25 ml added to cuvette).

The results of 2,3-DPG assays are best expressed in terms of Hb content or red cell volume. Thus, if the result of the previous calculation is represented by D, then:

or

and

where Hb = Hb in g/l of whole blood and 64 is the molecular weight of Hb × 10^–3.

The molar ratio of 2,3-DPG to Hb in normal blood is about 0.75:1.

Normal Range

The normal range is 4.5–5.1 µmol/ml packed red cells or 10.5–16.2 µmol/g Hb. Neonatal values are about 20% lower than adult.³⁹

Each laboratory should determine its own normal range.

Significance of 2,3-DPG Concentration

An increase in 2,3-DPG concentration is found in most conditions in which the arterial blood is undersaturated with oxygen, as in congenital heart and chronic lung diseases, in most acquired anaemias, at high altitudes, in alkalosis and in hyperphosphataemia. Decreased 2,3-DPG levels occur in hypophosphataemic states and in acidosis.

Acidosis, which shifts the oxygen dissociation curve to the right, causes a fall in 2,3-DPG, so that the oxygen dissociation curve of whole blood from patients with chronic acidosis (such as patients in diabetic coma or precoma) may have nearly normal dissociation curves. A rapid correction of the acidosis will lead to a major shift of the curve to the left (i.e. to a marked increase in the affinity of Hb for oxygen, which may lead to tissue hypoxia). Caution should therefore be exercised in correcting acidosis. Measurement of oxygen dissociation is described below.

From the diagnostic point of view, the main importance of 2,3-DPG determination is (1) in haemolytic anaemias and (2) in the interpretation of changes in the oxygen affinity of blood.

2,3-DPG levels are generally slightly lower than normal in HS and this probably accounts for the slight erythrocytosis that is sometimes seen after splenectomy. Extremely low red cell 2,3-DPG concentration associated with erythrocytosis has been reported in a kindred with complete 2,3-diphosphoglycerate mutase deficiency.⁵⁷

Interpretation

Figure 12.6 shows the sigmoid nature of the oxygen dissociation curve of Hb A and the effect of hydrogen ions on the position of the curve. A shift of the curve to the right indicates decreased affinity of the Hb for oxygen and hence an increased tendency to give up oxygen to the tissues; a shift to the left indicates increased affinity and so an increased tendency for Hb to take up and retain oxygen. Hydrogen ions, 2,3-DPG and some other organic phosphates such as ATP shift the curve to the right. The amount by which the curve is shifted may be expressed by the p₅₀O₂ (i.e. the partial pressure of oxygen at which the Hb is 50% saturated).

Figure 12.6 Effect of pH on the oxygen dissociation curve.

The oxygen affinity, as represented by the p₅₀O₂, is related to compensation in haemolytic anaemias;⁶² 1 g of Hb can carry about 1.34 ml of O₂. Figure 12.7 shows the O₂ dissociation curves of Hb A and Hb S plotted according to the volume of oxygen contained in 1 litre of blood when the Hb concentrations is 146 g/l and 80 g/l, respectively. The p₅₀O₂ of Hb A is given as 26.5 mmHg (3.5 kPa) and Hb S as 36.5 mmHg (4.8 kPa). It will be seen that in the change from arterial to venous saturation, the same volume of oxygen is given up despite the difference in Hb concentration. Patients with a high p₅₀O₂ achieve a stable Hb at a lower level than normal and this should be taken into account when planning transfusion for these patients.

Figure 12.7 Effect of O₂ affinity on O₂ delivery to tissues.

Bohr Effect

An increase in CO₂ concentration produces a shift to the right (i.e. a decrease in oxygen affinity). This effect originally described by C. Bohr,⁶³ is mainly a result of changes in pH, although CO₂ itself has some direct effect. The Bohr effect is given a numeric value, log p₅₀O₂/pH, where log p₅₀O₂ is the change in p₅₀O₂ produced by a change in pH (pH). The normal value of the Bohr effect at physiological pH and temperature is about 0.45.

Hill’s Constant (‘n’)

Hill’s constant (‘n’) represents the number of molecules of oxygen that combine with one molecule of Hb.⁶⁴ Experiments showed that the value was 2.6 rather than the expected 4. The explanation for this lies in the effect of binding 1 molecule of oxygen by Hb on the affinity for binding further oxygen molecules by Hb, the so-called allosteric effect of haem–haem interaction: ‘n’ is a measure of this effect and the calculation of the ‘n’ value helps in identifying abnormal Hbs, the molecular abnormality of which leads to abnormal haem–haem interaction.⁶⁵